Biocatalytic microcapsules for catalyzing gas conversion

ABSTRACT

Methods of forming such microcapsules, in accordance with some embodiments, include: emulsifying at least one biocatalyst in a polymer precursor mixture; emulsifying the polymer precursor mixture in an aqueous carrier solution; crosslinking one or more polymer precursors of the polymer precursor mixture to form a plurality of microcapsules each independently comprising: a polymeric shell permeable to one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell. In further embodiments, corresponding methods of using the inventive microcapsules for catalyzing one or more target gases using include: exposing a plurality of the biocatalytic microcapsules to the one or more target gases.

RELATED APPLICATIONS

This application is a divisional of U.S. patent application Ser. No. 15/411,337 filed Jan. 20, 2017, which is a continuation-in-part of U.S. patent application Ser. No. 15/003,257 filed Jan. 21, 2016, from all of which priority is claimed and which are all herein incorporated by reference.

The United States Government has rights in this invention pursuant to Contract No. DE-AC52-07NA27344 between the United States Department of Energy and Lawrence Livermore National Security, LLC for the operation of Lawrence Livermore National Laboratory.

FIELD OF THE INVENTION

The present invention relates to bioreactors, and more particularly to biocatalytic microcapsules providing improved surface area and mass transport to facilitate conversion of target gases using biocatalyst(s) in the biocatalytic microcapsules.

BACKGROUND

Most chemical reactions of interest for clean energy are routinely carried out in nature. These reactions include the conversion of sunlight to chemical energy, the transfer of carbon dioxide into and out of solution, the selective oxidation of hydrocarbons (including methane to methanol), the formation of C—C bonds (including methane to ethylene), and the formation and dissolution of Si—O bonds (including enhanced mineral weathering). Conventional industrial approaches to catalyze these reactions are either inefficient or have yet to be developed.

Certain enzymes have been identified that carry out each of the aforementioned reactions. Unfortunately, industrial biocatalysis is primarily limited to the synthesis of low-volume, high-value products, such as pharmaceuticals, due to narrow operating parameters required to preserve biocatalyst activity.

Conventional gas conversion involves enzyme-catalyzed reactions typically carried out in fermenters, which are closed, stirred, tank reactors configured to use bubbled gases for mass transfer. FIG. 1 , illustrates a conventional stirred tank reactor 100, which may include a motor 102, an input/feed tube 104, a cooling jacket 106, one or more baffles 108, an agitator 110, one or more gas spargers 112, and an aqueous medium 114. Gas exchange in the stirred tank reactor 100 may be achieved by bubbling from the sparger(s) 112 at the bottom of the aqueous medium 114 and gas collection above said aqueous medium 114. Care must be taken to maintain a narrow set of conditions in such stirred tank reactors to favor the desired metabolic pathways and discourage competing pathways and competing organisms. Moreover, stirred tank reactors are energy inefficient, require batch processing, suffer from loss of catalytic activity due to enzyme inactivation, and exhibit slow rates of throughput due to low catalyst loading and limited mass-transfer resulting from low solubility of the target gases in the liquid medium and relatively low surface area of the bubbled gases. Although surface area may be improved with increased stirring/agitation, this exacerbates the energy inefficiency of the stirred tank reactor and is an impractical solution to improving gas conversion efficiency.

To allow reuse of enzymes in stirred-tank reactors, and to improve stability in reactor conditions, enzymes may be immobilized on inert, artificial materials. As shown in FIG. 2 , one conventional approach is to immobilize enzymes 202 on the surface of an inert material 204. Other conventional approaches may involve immobilizing enzymes on the surface of accessible pores of inert materials. However, such conventional enzyme immobilization techniques also suffer from lower volumetric catalyst densities, low throughput rates, and do not have routes for efficient gas delivery or product removal. Accordingly, it would be advantageous to provide systems and techniques enabling the re-use of catalytic components while also providing high surface area and mass transport to improve the catalyst density, throughput, and efficiency of gas conversion.

In addition, certain applications in which gas conversion is important, e.g. preservation of organic or gas-sensitive materials such as food, medicine, etc. may prohibit the use of a liquid medium to facilitate mass transport. Accordingly, it would be a further advantage to provide systems and techniques enabling efficient gas conversion using dry compositions capable of effective mass transport and catalysis to preserve the sensitive material.

SUMMARY

In accordance with one embodiment, a method of forming microcapsules for selective catalysis of gases includes: emulsifying at least one biocatalyst in a polymer precursor mixture; emulsifying the polymer precursor mixture in an aqueous carrier solution; crosslinking one or more polymer precursors of the polymer precursor mixture to form a plurality of microcapsules each independently comprising: a polymeric shell permeable to one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell.

In further embodiments, a method for catalyzing one or more target gases using biocatalytic microcapsules includes: exposing a plurality of the biocatalytic microcapsules to the one or more target gases.

Other aspects and embodiments of the present invention will become apparent from the following detailed description, which, when taken in conjunction with the drawings, illustrate by way of example the principles of the invention.

BRIEF DESCRIPTION OF THE DRAWINGS

For a fuller understanding of the nature and advantages of the present invention, reference should be made to the following detailed description read in conjunction with the accompanying drawings.

FIG. 1 is a schematic representation of a conventional stirred tank reactor, according to the prior art.

FIG. 2 is a schematic representation of enzymes immobilized on an exterior surface of an inert material, according to the prior art.

FIG. 3 is a schematic representation of enzymatic reactive components embedded within a polymeric network, according to one embodiment.

FIG. 4 is a process flow illustrating a method for embedding enzymatic reactive components within a two phase (AB) polymer network, according to one embodiment.

FIG. 5 is a process flow illustrating a method for embedding enzymatic reactive components within a two phase (AB) polymer network, according to another embodiment.

FIG. 6 is schematic representation of a bioreactor comprising a hollow tube network/lattice configured to optimize mass transfer, according to one embodiment.

FIG. 7 is a flowchart of a method for forming a bioreactor via 3D printing, according to one embodiment.

FIG. 8 illustrates various photographs of PEG-pMMO 3D structures formed/patterned according to a projection microstereolithography (PμSL) process, according to some embodiments.

FIG. 9 is a process flow illustrating a method for forming a polyethylene glycol diacrylate (PEGDA) hydrogel comprising particulate methane monooxygenasae (pMMO), according to one embodiment.

FIG. 10A is a plot illustrating pMMO retention by weight in a PEGDA hydrogel as a function of the volume percentage of PEGDA present during polymerization, where 150 μg of pMMO is initially included within the PEGDA hydrogel.

FIG. 10B is a plot illustrating pMMO activity in a PEGDA hydrogel as a function of the volume percentage of PEGDA present during polymerization, where 150 μg of pMMO is initially included within the PEGDA hydrogel.

FIG. 10C is a plot illustrating pMMO retention by weight in a PEGDA hydrogel as a function of the amount of pMMO (μg) included during polymerization.

FIG. 10D is a plot illustrating the activity of PEGDA-pMMO and a pMMO control as a function of the amount of pMMO (μg) included during the activity assay.

FIG. 11A is a plot illustrating the activity of the PEGDA-pMMO hydrogel after reusing said hydrogel over multiple cycles.

FIG. 11B is a plot illustrating the amount of methanol (nmoles) produced per mg of pMMO for both as-isolated membrane bound pMMO and PEGDA-pMMO over twenty cycles of methane activity assay.

FIG. 12A is a schematic representation of a continuous flow-through PEGDA-pMMO hydrogel bioreactor, according to one embodiment.

FIG. 12B is a plot illustrating the amount of methanol (nmole) produced per mg of pMMO in the PEGDA-pMMO hydrogel bioreactor of FIG. 12A.

FIG. 13 is a plot illustrating the dependence of PEGDA-pMMO activity on surface area to volume ratio for a PEGDA-pMMO hydrogel bioreactor.

FIG. 14A is a simplified schematic of a biocatalytic microcapsule in accordance with one embodiment of the present disclosures.

FIG. 14B is a photographic representation of a plurality of biocatalytic microcapsules, according to another embodiment.

FIG. 15 depicts a simplified schematic of a microcapsule assembly apparatus according to one approach.

FIG. 16 is a flowchart of a method for forming biocatalytic microcapsules for selective catalysis of target gases, according to one embodiment.

FIG. 17 is a flowchart of a method for catalyzing one or more target gases using biocatalytic microcapsules, according to one embodiment.

DETAILED DESCRIPTION

The following description is made for the purpose of illustrating the general principles of the present invention and is not meant to limit the inventive concepts claimed herein. Further, particular features described herein can be used in combination with other described features in each of the various possible combinations and permutations.

Unless otherwise specifically defined herein, all terms are to be given their broadest possible interpretation including meanings implied from the specification as well as meanings understood by those skilled in the art and/or as defined in dictionaries, treatises, etc.

It must also be noted that, as used in the specification and the appended claims, the singular forms “a,” “an” and “the” include plural referents unless otherwise specified.

As also used herein, the term “about” when combined with a value refers to plus and minus 10% of the reference value. For example, a length of about 100 nm refers to a length of 100 nm±10 nm.

As further used herein, the term “fluid” may refer to a liquid or a gas.

As discussed previously, enzymes have been identified that catalyze virtually all of the reactions relevant to clean energy, such as selective transformations among carbon fuels, gas to liquids technology, storage of solar energy, exchange of CO₂, formation and dissolution of silicates, and neutralization of wastes. However, industrial enzyme biocatalysis is currently limited to low-volume, high-value products such as pharmaceuticals due to the narrow operating parameters required to preserve biocatalyst activity; slow rates of throughput due to low catalyst loading; limited mass transfer; and susceptibility to contamination and poisoning. These limitations require that many biocatalysis processes are carried out in single phase, aqueous media such as that provided in stirred tank reactors. However, stirred tank reactors are energy inefficient, require batch processing, and have poor mass transfer characteristics. While techniques have emerged to improve the stability and allow reuse of enzymes in stirred tank reactors, such techniques involve immobilizing the enzymes solely on the exterior surface(s) of an inert material or on the exterior surface(s) of the pores of an inert material. Unfortunately, these conventional immobilization techniques still fail to rectify the slow throughput rates and limited mass transfer associated with current biocatalysis processes.

To overcome the aforementioned drawbacks, embodiments disclosed herein are directed to a novel class of bioreactor that includes a membrane comprising one or more types of reactive enzymes and/or enzyme-containing cell fragments embedded within, and throughout the depth of, a multicomponent polymer network. In various approaches, this multicomponent polymer network may comprise two or more polymer types, or a mixture of a polymer and inorganic material. Preferably, the membrane comprises permeable, multi-component polymers that serve as both a mechanical support for the embedded enzymes, as well as functional materials configured to perform one or more additional functions of the bioreactor, such as: efficiently distributing reactants and removing products; exposing the embedded enzymes to high concentrations of reactants; separating reactants and products; forming high surface area structures for exposing the embedded enzymes to reactants; supplying electrons in hybrid enzyme-electrochemical reactions; consolidating enzymes with co-enzymes in nanoscale subdomains for chained reactions, etc. In additional approaches, this membrane may be molded into shapes/features/structures (e.g., hollow fibers, micro-capsules, hollow tube lattices, spiral wound sheets, etc.) to optimize the bioreactor geometry for mass transfer, product removal, and continuous processing.

The novel class of bioreactor disclosed herein may be especially suited to catalyze reactions that occur at phase boundaries, e.g., gas to liquid, liquid to gas, polar to non-polar, non-aqueous to aqueous, etc. Accordingly, the novel class of bioreactors disclosed herein may be useful for reactions in clean energy applications that involve a gas-phase reactant or product (e.g., methane to methanol conversion, CO₂ absorption, oxidation reactions with O₂, reduction reactions with H₂ or methane, CO₂ conversion to synthetic fuel, etc.), as well as reactions in the chemical and pharmaceutical industries that involve treatment of non-polar organic compounds with polar reactants (or vice versa).

The following description discloses several general, specific, and preferred embodiments relating to biocatalytic microcapsules providing improved surface area and mass transport to facilitate conversion of target gases using biocatalyst(s) in the biocatalytic microcapsules.

According to one general embodiment, a microcapsule for selective catalysis of gases, the microcapsule comprising: a polymeric shell permeable to one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell.

In accordance with another general embodiment, a method of forming microcapsules for selective catalysis of gases includes: emulsifying at least one biocatalyst in a polymer precursor mixture; emulsifying the polymer precursor mixture in an aqueous carrier solution; crosslinking one or more polymer precursors of the polymer precursor mixture to form a plurality of microcapsules each independently comprising: a polymeric shell permeable to one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell.

In further general embodiments, a method for catalyzing one or more target gases using biocatalytic microcapsules includes: exposing a plurality of the biocatalytic microcapsules to the one or more target gases.

Block Polymer/Copolymer Embodiments

Referring now to FIG. 3 , a membrane 300 particularly suitable for use in a bioreactor is shown according to one embodiment. As an option, the membrane 300 may be implemented in conjunction with features from any other embodiment listed herein, such as those described with reference to the other FIGS. Of course, the membrane 300 and others presented herein may be used in various applications and/or in permutations which may or may not be specifically described in the illustrative embodiments listed herein. For instance, the membrane 300 may be used in any desired environment and/or include more or less features, layers, etc. than those specifically described in FIG. 3 .

As shown in FIG. 3 , the membrane 300 includes a plurality of enzymatic reactive components 302 embedded within a polymer network 304. In various approaches, the enzymatic reactive components 302 may comprise about 1% to 80% of the mass of the polymer network 304. The enzymatic reactive components 302 may be configured to catalyze any of the reactions described herein, and in particular reactions that take place at phase boundaries (e.g., gas to liquid, liquid to gas, polar to non-polar, non-aqueous to aqueous, etc.).

In some approaches, the plurality of enzymatic reactive components 302 may comprise one or more of: isolated enzymes, trans-cell-membrane enzymes, cell-membrane-bound enzymes, liposomes coupled to/comprising an enzyme, etc. Stated another way, each enzymatic reactive component 302 may individually be selected from the group selected from: an isolated enzyme, a trans-cell-membrane enzyme, a cell-membrane-bound enzyme, and a liposome coupled to/comprising an enzyme. Suitable enzymatic reactive components 302 may include, but are not limited to, formate dehydrogenase, carbonic anhydrase, cytochrome p450, hydrogenase, particulate methane monooxygenasae (pMMO), photosynthetic complexes, etc. Moreover, while each of the enzymatic reactive components 302 may be the same (e.g., comprise the same structure and/or composition) in particular approaches; other approaches may require at least two of the enzymatic reactive components 302 to be different (e.g., have a different structure and/or composition) from one another.

In approaches where at least one of the enzymatic reactive components 302 includes a membrane-bound enzyme, said enzyme may be stabilized prior to incorporation into the polymer network 304. For instance, in one stabilization approach, cell fragments comprising the enzyme of interest may be used, and directly incorporated into the polymer network 304. In another stabilization approach, a lipopolymer may first be formed by linking a lipid to a polymer of interest. The lipid region of the polymer may spontaneously insert into the cell membrane, thereby creating a polymer functionalized liposome, which may be incorporated in the polymer network 304. In yet another stabilization approach, the enzyme of interest may be coupled to and/or encapsulated into a nano-lipo-protein particle (NLP), which may then be incorporated in the polymer network 304.

The enzymatic reactive components 302 may be incorporated into the polymeric network 304 via several methods including, but not limited to: attaching the enzymatic reactive components 302 to electrospun fibers of a first polymer, and backfilling with a second polymer (see, e.g., the method described in FIG. 4 ); directly incorporating the enzymatic reactive component 302 into a polymer or block-copolymer network before or after crosslinking the network (see, e.g., the method described in FIG. 5 ); and other suitable incorporation methods as would become apparent to one having skill in the art upon reading the present disclosure.

With continued reference to FIG. 3 , the polymeric network 304 may include at least a two phase polymer network, e.g. a polymer network comprising two or more polymeric materials. This polymer network 304 may be configured to serve as a mechanical support for the enzymatic reactive components 302 embedded therein, concentrate reactants, and remove products. In preferred approaches, the polymeric network 304 may include nanometer scale domains of higher reactant permeability, as well as nanometer scale domains of higher product permeability.

In particular approaches involving gas to liquid reactions, the polymeric network may include nanometer scale domains of higher gas permeability, such as silicon, as well as nanometer scale domains of higher product permeability, such as a polyethylene glycol (PEG) based hydrogel. These domains of high gas permeability typically also have higher gas solubility, increasing the local concentration of reactants (e.g., relative to the aqueous medium in a stirred tank reactor) and therefore increase the turnover frequency of the enzymatic reactive components 302; whereas, the domains of low gas permeability and high product permeability may efficiently remove the product and reduce product inhibition (thereby also increasing the turnover frequency and stability of the enzymatic reactive components 302) or serve to stabilize the enzymatic reactive components. In various approaches, the permeability for the “higher gas permeability phase” may be greater than 100 barrer.

In some approaches, the polymer network 304 may comprises a di-block copolymer network. In other approaches, the polymer network 304 may include a tri-block copolymer network. Suitable polymers for the polymeric network 304 may include silicone polymers, polydimethylsiloxane (PDMS), poly(2-methyl-2-oxazoline) (PMOXA), polyimide, PEG, polyethylene glycol diacrylate (PEGDA), poly(lactic acid) (PLA), polyvinyl alcohol (PVA), and other such polymers compatible with membrane proteins and block copolymer synthesis as would become apparent to one skilled in the art upon reading the present disclosure. In more approaches, each polymer in the polymeric network 304 may have a molecular weight ranging from about 500 Daltons to about 500 kiloDaltons.

In other approaches, the polymeric network 304 may include a mixture of at least one polymer material and at least one inorganic material.

In various approaches, a thickness, t₁, of the enzyme embedded polymer network 304 may be in a range from about 1 micrometer to about 2 millimeters.

As indicated above, the membrane 300 may be configured to separate the reactants and products associated with a catalyzed reaction of interest. These reactants and products may be two different fluids, such as liquids and gasses, aqueous species and non-aqueous species, polar species and non-polar species, etc. In one exemplary approach where the membrane 300 may be configured to separate methane and oxygen from methanol, the methane reactant concentration may be in a range from about 1 to about 100 mM, the oxygen reactant concentration may be in a range from about 1 to about 100 mL, and the methanol product concentration range may be in a range from about 0.1 to about 1000 mM.

To further facilitate reactant-production separation, at least a portion of one surface of the membrane 300 may include an optional reactant permeable polymer layer 306 coupled thereto, as shown in FIG. 3 . In preferred approaches, this reactant permeable polymer layer 306 may also be impermeable to products generated from the reactions catalyzed by the enzymatic reactive components 302. Suitable polymeric materials for this reactant permeable polymer layer 306 may include, but are not limited to, nanofiltration, reverse-osmosis, or chemically selective membranes, such as poly(ethylene imine), PVA, poly(ether ether ketone) (PEEK), cellulose acetate, or polypropylene (PP). In some approaches, a thickness, t₂, of the reactant permeable polymer layer 306 may be in a range from about 0.1 to about 50 micrometers. This optional reactant permeable polymer layer 306 may be particularly suited for approaches involving an organic polar reactant and an organic non-polar product (and vice versa).

As also shown in FIG. 3 , at least a portion of one surface of the membrane 300 may include an optional product permeable polymer layer 308 coupled thereto. This product permeable polymer layer 308 may preferably be coupled to a surface of the membrane 300 opposite that on which the reactant permeable polymer layer 306 is coupled, thereby facilitating entry of reactants (e.g., gaseous reactants) on one side of the membrane 300, and removal of the reaction products (e.g., liquid reaction products) on the opposing side of the membrane 300. In more preferred approaches, this product permeable polymer layer 308 may also be impermeable to the reactants introduced into the enzyme embedded polymer network 304. Suitable polymeric materials for this product permeable polymer layer 308 may include, but are not limited to, nanofiltration, reverse-osmosis, or chemically selective membranes, such as poly(ethylene imine), PVA, poly(ether ether ketone) (PEEK), cellulose acetate, or polypropylene (PP). In some approaches, a thickness, t₃, of the product permeable polymer layer 308 may be in a range from about 0.1 to about 50 micrometers.

In some approaches, one or more of the enzymatic reactive components 302 may require a cofactor for function. Accordingly, cofactors may be supplied by co-localized enzymes in reactor domains of the polymer network 304 (not shown in FIG. 3 ), and/or be retained within a cofactor impermeable layer coupled to a portion of the membrane 300 (not shown in FIG. 3 ).

In various approaches, a total thickness, t₄, of the membrane 300 may be in a range from about 10 to about 3100 micrometers.

In yet more approaches, the membrane 300 may be shaped into features, structures, configurations, etc. that provide a desired surface area to support efficient transport of reactants to, and products from, the enzymatic reactive components 302. For instance, the membrane 300 may be shaped into at least one of: a hollow fiber membrane, a micro-capsule membrane, a hollow tube membrane, a spiral wound membrane, etc.

Referring now to FIG. 4 , a method 400 for embedding enzymatic reactive components within a two phase (AB) polymer network is shown according to one embodiment. As an option, the present method 400 may be implemented in conjunction with features from any other embodiment listed herein, such as those described with reference to the other FIGS. Of course, this method 400 and others presented herein may be used to form structures for a wide variety of devices and/or purposes, which may or may not be related to the illustrative embodiments listed herein. It should be noted that the method 400 may include more or less steps than those described and/or illustrated in FIGS. 4 , according to various embodiments. It should also be noted that that the method 500 may be carried out in any desired environment.

As shown in FIG. 4 , an enzymatic reactive component 402 is adsorbed to at least one portion of the exterior surface of polymer A 404, thereby forming enzyme-embedded polymer A 406. In preferred approaches, polymer A 404 may comprise one or more hydrophobic, reactant permeable (e.g., gas permeable) polymeric materials configured to provide high concentrations and fast transport of reactants. In further approaches, polymer A 404 may be a polymer nanofiber generated using electrospinning, extrusion, self-assembly, or other suitable technique as would become apparent to one skilled in the art upon reading the present disclosure. In additional approaches, such a polymer A nanofiber may be crosslinked to other polymer A nanofibers. In one exemplary approach, polymer A 404 comprises PDMS.

In various approaches, the enzymatic reactive component 402 may be selected from the group consisting of: an isolated enzyme, an enzyme comprising a cell fragment (e.g., a cell membrane or cell membrane fragment), and a liposome comprising/coupled to an enzyme. In some approaches, the enzymatic reactive component 402 may include at least one of: formate dehydrogenase, carbonic anhydrase, cytochrome p450, hydrogenase, particulate methane monooxygenasae (pMMO), photosynthetic complexes, etc.

In the non-limiting embodiment shown in FIG. 4 , a plurality of enzymatic reactive components 402 may be adsorbed to one or more portions of the exterior surface of polymer A 404. These enzymatic reactive components 402 may be adsorbed to at least the majority, or more preferably about an entirety, of the exterior surface of polymer A 404. The lipid bilayer vesicles of the enzymatic reactive components 402 may spontaneously collapse on the exterior surface of polymer A 404, thereby forming a lipid-bilayer functionalized surface.

As further shown in FIG. 4 , the enzyme-embedded polymer A 406 may be mixed with polymer B 408 to create the two phase (AB) polymer monolith 410 with the enzymatic reactive components 402 at the interface between the two phases. In preferred approaches, polymer B 408 may comprise one or more hydrophilic, product permeable polymeric materials configured to provide transport of products, as well as stabilize the enzymatic reactive components 402. For instance, in one specific approach, polymer B 408 may be a hydrophobic polymer hydrogel.

While the resulting polymeric network shown in FIG. 4 includes two phases (i.e., polymer A and polymer B), it is important to note that said polymeric network may include more than two phases in additional approaches.

Referring now to FIG. 5 , a method 500 for embedding enzyme reactive components within a two phase (AB) polymer network is shown according to another embodiment. As an option, the present method 500 may be implemented in conjunction with features from any other embodiment listed herein, such as those described with reference to the other FIGS. Of course, this method 500 and others presented herein may be used to form structures for a wide variety of devices and/or purposes, which may or may not be related to the illustrative embodiments listed herein. It should be noted that the method 500 may include more or less steps than those described and/or illustrated in FIGS. 5 , according to various embodiments. It should also be noted that that the method 500 may be carried out in any desired environment.

As shown in FIG. 5 , enzymatic reactive components 502 may be directly incorporated in a block copolymer network 504 prior to or after cross-linking said network. As described herein, each enzymatic reactive component 502 may be independently selected from an isolated enzyme, an enzyme comprising a cell fragment (e.g., a cell membrane or cell membrane fragment), and a liposome comprising/coupled to an enzyme. In some approaches, the enzymatic reactive component 502 may include at least one of: formate dehydrogenase, carbonic anhydrase, cytochrome p450, hydrogenase, particulate methane monooxygenasae (pMMO), photosynthetic complexes, etc.

As shown in the non-limiting embodiment of FIG. 5 , the block copolymer network 504 is a di-block copolymer network comprising two different polymers (polymer A 506 and polymer B 508). In preferred approaches, polymer A 506 may comprise one or more reactant permeable, hydrophobic polymeric materials, whereas polymer B 508 may comprise one or more product permeable, hydrophilic polymeric materials. It is again important to note that while the block copolymer network 504 shown in FIG. 5 includes two phases (i.e., polymer A 506 and polymer B 508), said block copolymer network may include more than two phases in other approaches.

In various approaches, the enzymatic reactive components 502 may be incorporated directly into the block copolymer network 504 using lipopolymers (preferably di-block lipopolymers). Lipopolymers may be generated by linking a lipid to a polymer of interest, such as PEG, creating PEG-lipid conjugates, such as PEG-phosphatidylethanolamie. The lipid region of the polymer may spontaneously insert into the cell membrane, thereby creating a polymer functionalized liposome.

Referring now to FIG. 6 , a bioreactor 600 comprising a network/lattice of three dimensional structures configured to optimize mass transfer is shown according to one embodiment. As an option, the bioreactor 600 may be implemented in conjunction with features from any other embodiment listed herein, such as those described with reference to the other FIGS. Of course, the bioreactor 600 and others presented herein may be used in various applications and/or in permutations which may or may not be specifically described in the illustrative embodiments listed herein. For instance, the bioreactor 600 may be used in any desired environment and/or include more or less features, layers, etc. than those specifically described in FIG. 6 .

As noted above, the bioreactor 600 includes a network/lattice 602 of three dimensional structures. As particularly shown in FIG. 6 , the network/lattice 602 includes multiple layers (e.g., 2, 3, 4, 5, 6, 7, or more layers, etc.) of three-dimensional (3D) hollow tubes 604. It is important to note, however, that the hollow tube network/lattice 602 of the bioreactor 600, and others disclosed herein, may include one or more layers of three-dimensional hollow tubes 604 in various approaches. The hollow tubes 604 may preferably be oriented in the lattice such that their hollow interiors are perpendicular to a thickness direction of the lattice (e.g., perpendicular to the z axis shown in FIG. 6 ).

In some approaches, the bioreactor 600 may have a thickness (as measured parallel to the z-axis in FIG. 6 ) in a range from about 1 to about 300 cm, and a length (as measured in a direction parallel to the y-axis of FIG. 6 ) and width (as measured in a direction parallel to the x-axis of FIG. 6 ) scaled to the application, ranging from about 2 cm for laboratory applications to 10 meters for industrial applications.

The walls of each hollow tube 604 may comprise a membrane material 606, such as the membrane material of FIG. 3 , configured to separate reactants (e.g., gaseous reactants) and products (e.g., hydrophilic products). Accordingly, the hollow tubes 604 form polymer microchannels through which the hydrophilic reaction products may flow.

As particularly shown in FIG. 6 , the membrane material 606 of each hollow tube 604 may comprise a plurality of enzymatic reactive components 608 (e.g., isolated enzymes, membrane-bound enzymes, liposomes comprising/couple to an enzyme, etc.) embedded throughout a polymer network 610. The polymer network 610 may comprise reactant permeable fibrils of a first polymer 612 that increase the local concentration of reactants and enhance mass transfer throughout the membrane material 606. In some approaches, the enzymatic reactive components 608 may be immobilized on the fibrils of the first polymer 612. The polymer network 610 may also include at least another polymer material (e.g., a hydrogel matrix material) configured to hydrate the enzymatic reactive components 608 and provide a route for hydrophilic product removal. The membrane material 606 may also include an optional reactant permeable (product impermeable) layer 614 coupled to one side (e.g., an exterior side) of the polymer network 610 and/or a product permeable (reactant impermeable) layer 616 coupled to the opposite side (e.g., an interior side) of the polymer network 610. The optional product permeable (reactant impermeable) layer 616 may also facilitate product removal and prevent coenzyme and/or cofactor diffusion into the liquid core that contains the desired products.

The thickness, tmem, of the membrane material 606 may be in a range from about 10 to about 1000 micrometers. In some approaches, t_(n), may be about 300 p.m. Additionally, The thickness, ttube, of each hollow tube 604 may be in a range from about 10 micrometers to about 10 millimeters. In various approaches, ttube may be about 1 mm. In yet more approaches, the length, l_(tube), of each hollow tube 604 may be in a range from about 5 centimeters to about 10 meters.

It is important to note that while the cross section of each hollow tube 604, as taken perpendicular to the y-axis of FIG. 6 , is shown a circular, this need not be the case. For instance, in other approaches, each hollow tube 604 may have a cross sectional shape that is elliptical, rectangular, square, triangular, irregular shaped, etc. Moreover, in preferred approaches, each hollow tube 604 may have the same cross sectional shape, materials, and/or dimensions; however, this again need not be case. For instance, in alternative approaches, at least one of the hollow tubes 604 may have a cross sectional shape, materials, and/or dimensions that are different than that of another of the hollow tubes 604.

In one particular approach, one or more of the hollow tubes 604 in at least one of the layers may differ from one or more hollow tubes 604 in at least another of the layers with respect to: cross sectional shape, and/or one or more membrane material(s), and/or one or more dimensions. In another particular approach, one or more of the hollow tubes 604 in at least one of the layers may differ from at least another hollow tube 604 in the same layer with respect to: cross sectional shape, and/or one or more membrane materials, and/or one or more dimensions.

In yet further approaches, the spacing between the hollow tubes 604 in at least one of the layers may be about uniform. In more approaches, the spacing between the hollow tubes 604 in at least one of the layers may vary throughout the layer. For example, in one such approach, at least one of the layers may have at least one area having an average spacing, s₁, between adjacent hollow tubes 604, and at least a second area having an average spacing s₂, where s₁ and s₂ are different. In yet other approaches, the spacing between the hollow tubes 604 in at least one of the layers may differ from the spacing between the hollow tubes 604 of at least another of the layers.

Referring now to method 7, an exemplary method 700 of forming a bioreactor (such as those disclosed herein) is shown, according to one embodiment. As an option, the present method 700 may be implemented in conjunction with features from any other embodiment listed herein, such as those described with reference to the other FIGS. Of course, the method 700 and others presented herein may be used in various applications and/or in permutations, which may or may not be specifically described in the illustrative embodiments listed herein. Moreover, more or less operations than those shown in FIG. 7 may be included in method 700, according to various embodiments. Furthermore, while exemplary processing techniques are presented with respect to FIG. 7 , other known processing techniques may be used for various steps.

As shown in FIG. 7 , the method 700 includes forming a membrane material comprising: a polymeric network configured to separate a first fluid from a second fluid, and a plurality of enzymatic reactive components embedded/incorporated within the polymeric network. See operation 702.

The enzymatic reactive components may comprise any of the enzymatic reactive components disclosed herein including, but not limited to, isolated enzymes, trans-cell-membrane enzymes, cell-membrane-bound enzymes, liposomes coupled to/comprising an enzyme, combinations thereof, etc. Moreover, as discussed previously, the enzymatic reactive components may be embedded/incorporated into the polymeric network via several methods including, but not limited to: attaching the enzymatic reactive components to electrospun fibers of a first polymer, and backfilling with a second polymer (see, e.g., the method described in FIG. 4 ); directly incorporating the enzymatic reactive component into a polymer or block-copolymer network before or after crosslinking the network (see, e.g., the method described in FIG. 5 ); and other suitable incorporation methods as would become apparent to one having skill in the art upon reading the present disclosure.

The polymeric network may include any of the materials, and/or be of the same form, as any of the polymeric networks disclosed herein. For instance, this polymer network may be configured to serve as a mechanical support for the enzymatic reactive components embedded therein, as well as include nanometer scale domains of higher permeability to the first fluid and nanometer scale domains of higher permeability to the second fluid. Moreover, in some approaches, the polymeric network may include at least a two phase polymer network, e.g. a polymer network comprising two or more polymeric materials. In other approaches, the polymeric network may include a mixture of at least one polymer material and at least one inorganic material.

As indicated above, the polymeric network may be configured to separate a first and second fluid associated with a reaction catalyzed by the enzymatic reactive components embedded therein. The first and second fluids may be two different fluids, such as liquids and gasses, an aqueous species and a non-aqueous species, a polar species and a non-polar species, etc.

As also shown in FIG. 7 , the method 700 includes fabricating and patterning one or more layers in the membrane material via a 3D printing process. See operation 704. In preferred approaches, the 3D printing process includes a_projection microstereolithography (PμSL) process as known in the art. In various approaches, each layer in the membrane material patterned/formed via the desired 3D printing process may include a plurality of three dimensional structures (e.g., hollow fibers, micro-capsules, hollow tube lattices, spiral wound sheets, etc.) configured to optimize the bioreactor geometry (and surface area) for mass transfer, reaction rate, product removal, continuous processing, etc. Photographs of several exemplary PEG-pMMO 3D structures formed/patterned according to a PμSL process are shown in FIG. 8 .

As discussed in greater detail below, the novel bioreactors described herein, such as described in FIG. 6 , may be particularly configured for methane activation with an energy efficiency from greater than or at least equal to about 68%. In such an approaches, the enzymatic reactive components embedded within the polymeric network may include pMMO to covert methane reactants, CH₄, to methanol products, CH3OH. Preferably, this engineered pMMO may exhibit a specific activity greater than about 5 μm/(gs) and/or a turnover frequency greater than about 10/s. Additionally, the amount of the engineered pMMO in such bioreactors may be about 50 g per L of reactor volume.

In some approaches, the aforementioned engineered pMMO may require a reducing agent for methane conversion. However, in other approaches, the engineered pMMO may not need such a reducing agent, or be configured to accept electrons via direct electron transfer. For instance, as shown in Table 1, the methane conversion may proceed by: (1) using pMMO configured to use methane as a reducing agent (reaction 1); (2) supplying electrons directly to the pMMO (reaction 2); and (3) using H2 gas. Yet another reaction pathway may involve steam reformation as shown in reaction 3.

TABLE 1 Energy Carbon Reaction Pathway efficiency efficiency Reaction 1: 2CH₄ + O₂ → 2CH₃OH 80% 100% Reaction 2: CH₄ + O₂ + 2H⁺ + 2e⁻ → >65%  100% CH₃OH + H₂O Reaction 3: 4CH₄ + 3O₂ → 3CH₃OH + 68%  75% CO + 2H₂O

Experiments/Examples

The following experiments and examples pertain to various non-limiting embodiments of the bioreactors described herein. In particular, the following experiments and examples are directed to bioreactors comprising pMMO embedded in a polymeric network for the conversion of methane to methanol. It is important to note that the following experiments and examples are for illustrative purposes only and do not limit the invention in anyway. It should also be understood that variations and modifications of these experiments and examples may be made by those skilled in the art. without departing from the spirit and scope of the invention

Overview

Advances in oil and gas extraction techniques have made vast new stores of natural gas (composed primarily of methane) available for use. However, substantial quantities of methane are leaked, vented, or flared during these operations. Indeed, methane emissions contribute about ⅓ of current net global warming potential. Compared to other hydrocarbons, and especially compared to the oil that is co-produced in hydrofracturing operations, methane has a much lower market value due to difficulty in methane storage and transport, and because methane has limited use as a transportation fuel.

Conversion of methane to methanol via conventional industrial technologies, such as steam reformation followed by the Fischer-Tropsch process, operate at high temperature and pressure, require a large number of unit operations, and yield a range of products. Consequently, conventional industrial technologies have a low efficiency of methane conversion to final products and can only operate economically at very large scales. There is thus a need in the art for a technology to efficiently convert methane to other hydrocarbons, and particularly to convert “stranded” sources of methane and natural gas (sources that are small, temporary, or not close to a pipeline) to liquids for later collection.

The only known true catalyst (industrial or biological) to convert methane to methanol under ambient conditions with 100% selectivity is the enzyme methane monooxygenase (MMO), found in methanotrophic bacteria, which converts methane to methanol according to the following reaction:

Partial methane oxidation by MMO enzymes can be carried out using whole methanotroph organisms, but this approach inevitably requires energy for upkeep and metabolism of the organisms, which reduces conversion efficiency. Moreover, biocatalysis using whole organisms is typically carried out in low-throughput unit operations, such as a stirred tank reactor.

One industrial-biological approach may therefor include separating the MMO enzyme from the host organism. Isolated enzymes may offer the promise of highly controlled reactions at ambient conditions with higher conversion efficiency and greater flexibility of reactor and process design. MMOs have been identified in both soluble MMO (sMMO) and particulate (pMMO) form. The use of pMMO has advantages for industrial applications because pMMO comprises an estimated 80% of the proteins in the cell membrane, Moreover, concentrating pMMO to a reasonable purity requires only isolating the membrane fraction of the lysed cells using centrifugation.

Traditional methods of enzyme immobilization and exposure to reactants are not sufficient to use pMMO effectively. These typical methods include cross-linking enzymes or immobilizing them on a solid support so that they can be separated from the products and carrying out batch reactions in the aqueous phase in a stirred tank reactor. As discussed previously, operation of a stirred tank reactor has several drawbacks, including low productivity, high operating costs, loss of catalytic activity due to enzyme inactivation, and variability in the quality of the product. The stirred-tank reactor is also not the optimal design for gas to liquid reactions such as methane to methanol conversion, as it does not allow efficient delivery of reactant gases to enzymes or organisms in the bulk solution. Gas delivery in stirred tank reactors is often achieved by bubbling the gas through the liquid, but this approach suffers from mass-transfer limitations. Furthermore, methane and oxygen are only sparingly soluble in aqueous solvents: 1.5 mM/atm and 1.3 mM/atm respectively at 25° C. Reactant concentrations are necessarily solubility-limited when the enzymes or organisms are dispersed in the aqueous phase.

Moreover, another reason as to why the pMMO enzyme is not amenable to standard immobilization techniques designed for soluble proteins is due to the fact that surfactant solubilization of isolated pMMO leads to a pronounced reduction in activity. For example, high surface area porous inorganic supports have been extensively studied and implemented for immobilizing soluble enzymes, and have been shown to enhance enzyme stability while achieving high enzyme loading in nanometer scale pores. The majority of the surface area in mesoporous materials is accessible only to proteins significantly smaller than 50 nm, and would therefore be inaccessible to the large (>100 nm), optically opaque vesicles and liposomes that comprise pMMO in crude membrane preparations.

Accordingly, the exemplary embodiments discussed in this experimental section are directed toward advances in biocatalytic processes for selective methane conversion. For instance, said exemplary embodiments are particularly directed toward a biocatalytic material comprising pMMO embedded in polyethylene glycol diacrylate (PEGDA) hydrogel. Embedding enzymes, such as pMMO that operate on gas phase reactants within the solid, gas permeable polymer hydrogel allows tuning of the gas solubility, permeability, and surface area thereof. An additional advantage to immobilizing pMMO within the polymer hydrogel, rather than on the surface of an impermeable support, is the potential to fully embed pMMO throughout the depth of the polymer hydrogel for high loading. PEGDA was selected as the primary polymer substrate because of its biocompatibility and flexibility for further development. PEGDA may be physically or chemically combined with hydrophobic polymers in additional approaches for enhanced gas solubility and transport in various approaches. Moreover, the pMMO embedded PEGDA hydrogel is amenable to various forms of 3D-printing, which offers the ability to rapidly prototype structures, tune micron to centimeter-scale material architecture, and precisely tailor structures for the system configuration and mass transfer, heat, and diffusion limitations.

Characterization of Block Copolymer Embodiments

a. pMMO activity in PEG hydrogel

Several methods for embedding pMMO in a PEGDA based polymer hydrogel were explored to enable its use as a biocatalytic material which could be molded into controlled, predetermined structures with tunable permeability and surface area for practical use. Initial efforts focused on solubilizing the crude membrane preparations using surfactant so that the material could be incorporated homogeneously in the polymer. It was discovered that any contact of the crude membrane preparations with surfactant, including encapsulation in nanolipoprotein particles, led to a pronounced decrease in activity. However, mixing the crude membrane fractions, either as prepared or extruded as liposomes directly with low concentrations of PEGDA 575 gave promising results. According the experiments described in this section focused on optimizing the activity and protein retention of crude membrane preparations with PEGDA 575.

A schematic of the method used to fabricate the PEG-pMMO hydrogels is shown in FIG. 9 . The synthesis of the PEG-pMMO materials required only membrane bound pMMO, PEGDA macromer, photoinitiator (not shown), and ultraviolet (UV) light. Photoinitiator concentrations higher than 0.5 vol % in PEGDA decreased the pMMO activity, therefore the photoinitiator concentration was held constant at 0.5 vol %.

Membrane bound pMMO alone in each activity assay as a positive control. The measured activity of the membrane bound pMMO alone was highly variable from experiment to experiment, from about 75 to 200 nmol MeOH mg⁻¹ min⁻¹, while the optimized PEG-pMMO samples were less variable, in a range from 65 to 128 nmol MeOH mg⁻¹ min⁻¹. The measured activity for both membrane bound pMMO alone and immobilized pMMO are similar to known values for membrane bound pMMO with methane as a substrate: 25-130 nmol MeOH mg⁻¹ min⁻¹.

FIGS. 10A-10D shows the results from systematically increasing the volume % of PEGDA in the solution prior to curing on protein retention (FIGS. 10A, 10C) and activity (FIGS. 10B, 10D). Mixing the pMMO solution with PEGDA at the appropriate vol % (10-80%), and UV curing resulted in 50 μl solid PEG-pMMO hydrogels. As the PEGDA vol % was increased from 10-80%, the overall stiffness of the material increased and the amount of residual liquid on the surface of the hydrogel decreased. A gradual increase was observed in the fraction of pMMO that was retained (0.4-0.75) when the PEGDA vol % was increased from 10-80% (FIG. 10A).

However, a dramatic decrease in pMMO activity was observed as the PEGDA vol % was increased (FIG. 10B). At 10% PEGDA, the pMMO activity was approximately 88+/−4 nmol MeOH min⁻¹mg⁻¹, which closely corresponded to the activity of pMMO alone (96+/−15 nmol MeOH min⁻¹mg⁻¹) (FIG. 10B). This value dropped below 30 nmol MeOH min⁻¹mg⁻¹ when the PEGDA vol % was greater than 50% (FIG. 10B). The amount of pMMO retained in the hydrogel before and after the activity assay did not change, indicating that no pMMO leached out during the activity assay and the enzyme was efficiently entrapped in the hydrogel. These combined findings demonstrate that one must consider both pMMO retention and activity when identifying the optimal PEGDA vol %. Since only a marginal increase in pMMO retention (0.4 vs 0.42) and a more significant decrease in pMMO activity (88 vs 74 nmol MeOH min⁻¹mg⁻¹) was observed when the PEGDA vol % was increased from 10% to 20%, all remaining experiments were performed using 10 vol % PEGDA.

FIGS. 10C and 10D illustrate the effect of varying the concentration of pMMO during hydrogel fabrication on pMMO retention and activity. For these experiments, the amount of pMMO used to generate the 50 μl PEG-pMMO hydrogel was varied between 50 μg and 550 μg. The fraction of pMMO retained was the highest at the lowest pMMO concentration tested (50 μg−0.75 retained) and a dramatic decrease was observed when the pMMO was increased to 150 μg (˜0.4 retained) (FIG. 10A). Further changes in the total pMMO retained was not observed when the pMMO was increased up to 550 μg. To assess the effect of varying the pMMO concentrations in the PEG-pMMO hydrogel on activity, PEG-pMMO hydrogels were prepared with 50-550 μg of pMMO, which resulted in retention of 35-200 μg of pMMO in the hydrogel, and the activity was measured. As shown in FIG. 10B, pMMO activity in the hydrogel was similar to the activity of pMMO alone when the amount of pMMO retained was below 50μg; however, there was a gradual decrease in pMMO activity in the hydrogels as the pMMO levels were increased from 50-200μg, which was not observed in the pMMO alone sample (FIG. 10D).

Preserving the native activity of pMMO in the PEG hydrogel required a balance between pMMO loading and enzyme activity. Higher polymer concentrations gave rise to higher pMMO loading and retention (FIG. 10A). Increasing the polymer concentration also correlated with diminished pMMO activity. This trend may be due to reduced polymer permeability or enzyme degradation by acrylate groups and/or free radicals at higher polymer concentrations. While it has been shown that PEDGA concentration (and by correlation, crosslinking density) has minimal effect on methane permeability in the gas phase, gas permeability is affected by the hydration (swelling) of hydrogel materials. Thus, PEGDA concentration may impact methane permeability in swollen PEG-pMMO. Higher PEGDA concentrations also decrease the distance between crosslinks and the diffusion of aqueous solutes through the hydrogel. Therefore, higher PEGDA concentrations may limit diffusion of the NADH cofactor to the enzyme or diffusion of the methanol product from the active site. Additionally, photo-initiated cross-linking reaction used to generate the cross-linked hydrogel results in the generation of free radicals, which can result in the oxidation of amino acids in proteins and cleavage of peptide bonds. The optimized PEG-pMMO formulations described in the text were remarkable in that they preserved physiological pMMO activity in a polymeric material; if a higher protein or polymer content is required the above issues might be managed by changing the macromer length and/or curing chemistry, in order to increase hydrogel mesh size (promoting diffusion) and reduce the number of radicals generated.

b. Reuse and Stability of PEG-pMMO Hydrogels

The development of fully active pMMO in a polymer material allowed the reuse of pMMO without painstaking centrifugation with each new set of reactants. Measurements were made regarding the effects of reuse of the PEG-pMMO hydrogel on overall enzyme activity and methanol generation using PEG-pMMO that was prepared with an initial pMMO amount of 150 μg and 10 vol % pMMO (FIGS. 11A-11B). In these experiments, the PEG-pMMO hydrogels were subjected to 20 cycles of 4 min exposures to methane. The hydrogel was washed thoroughly between each cycle to ensure that no residual methanol product remained in the hydrogel between cycles. The protein content in the reaction buffer for each cycle was measured to verify that the pMMO concentrations remained constant, and that there was no leaching through the course of the study. As shown in FIG. 11A, the activity between assay cycles 1 to 5 remained close to the initial activity (˜80 nmol MeOH min⁻¹mg⁻¹) and then gradually decreased to ˜45 nmol MeOH min⁻¹mg⁻¹ after 20 cycles. The error bars correspond to the standard deviation from the average of four replicates. FIG. 11B shows the cumulative methanol produced from these 20 consecutive reactions of PEG-pMMO compared to a single reaction of membrane bound pMMO. Immobilization of fully active pMMO in a material allowed the facile production of 10 fold more methanol per protein than could be produced with membrane bound pMMO (which can only be reused with painstaking repeated centrifugation and rinsing steps).

c. Continuous Flow-through Bioreactor

Establishing that that the PEG-pMMO material could be reused with no measureable protein leaching indicated that the material would be amenable for use in a bench-scale continuous flow reactor. A design where the pMMO material is suspended between gas and liquid reservoirs was discovered herein as desirable given that pMMO acts upon gas phase reactants and generates liquid phase. However, PEG-pMMO, and hydrogels in general, are mechanically brittle and difficult to handle when molded as thin membranes. Accordingly, the PEG-pMMO material was embedded into a three dimensional silicone lattice (printed using Direct Ink Write) in order to greatly increase the mechanical stability and to easily tune the size and shape of the hydrogel for use in a continuous reactor (FIG. 12A). As discussed in greater detail below, the lattice was constructed of 250 micron silicone struts and contained 250 micron void spaces (50% porosity) which were then infilled with PEGDA 575, crude pMMO membrane preparations, and photoinitiator and crosslinked in place with ultraviolet light. Two such lattice structures, thin and thick, were designed to compare effects of PEG-pMMO surface area to volume ratio on methanol production. The surface area to volume ratio of thin vs. thick for these experiments was 5 to 1. The silicone lattice structure increases the bulk gas permeability of the materials, since silicone permeability is at least 50 times greater than the PEGDA hydrogel permeability.

The resulting hybrid silicone-PEG-pMMO lattice materials were mechanically robust, allowing the suspension of the PEG-pMMO lattice of 1 millimeter thickness between gas and liquid reservoirs in a flow-through reactor. A schematic of the reactor cross section is shown in FIG. 12A. With this configuration, a methane/air gas mixture was flowed on one side of the lattice and the NADH was introduced on the other side, while continuously removing and collecting methanol in buffer. In order to determine the length of time the membrane could be continuously used, the cumulative methanol produced per mg of enzyme was measured at 25° C. at 30 min intervals in the thick lattice over the course of 5.5 hours. The methanol production rate (slope of methanol vs. time curve) was stable for about 2.5 hours, and declined gradually over the next 3 hours. In order to evaluate whether the geometry of PEG-pMMO material influenced methanol production rates, reactor outlet fractions from reactors containing the thin and thick lattices were compared at 15 min intervals at 45° C. over the course of two hours (FIG. 12B) in triplicate. The methanol concentrations produced in the flow reactor were on average 12 and 6% of what was predicted, for thin and thick lattices, respectively, based upon analyte flow rates and an assumed pMMO activity of 80 nmol MeOH min⁻¹mg⁻¹. The low concentration values relative to predicted values may be due to lower actual pMMO concentrations in the material than was calculated. As shown in FIG. 12B, the methanol produced (per mg of protein) by the thin membrane was double that produced by the thick membrane over the course of the first hour. Over the following hour, the methanol production rate by the thin membrane declined relative to that of the thick membrane; after two hours the average total methanol produced by the thin membrane was 1.5 times higher than that produced by the thick membrane. The results demonstrate that the ability to tune the geometry of immobilized pMMO, even at the millimeter scale, impacts the performance of the biocatalytic material.

d. Direct Printing of PEG-pMMO Hydrogels

Projection microstereolithography (PμSL) allows three dimensional printing of light-curable materials by projecting a series of images on the material, followed by changing the height of the stage at discrete increments, with micron-scale resolution in all three dimensions. Therefore, it was an ideal technique for directly printing the PEG-pMMO material and determining whether changing geometrical features of the material at these length scales can influence activity. PμSL was thus used to print PEG-pMMO lattice structures with increased surface area to volume ratio due to 100 μm² vertical channels corresponding to ˜15% void volume. In this experiment, the pMMO concentration of 5 mg/ml did not attenuate the light enough for highest resolution printing; consequently feature resolution was reduced in the z-direction and each layer of printed pMMO was exposed to multiple exposures to UV light. The pMMO activity in the printed cubic lattices with a total volume of about 27 mm³, which took approximately 50 min to print using PμSL, was reproducible but modest at 29 nmol MeOH min⁻¹mg⁻¹. The reduction in activity compared to crude pMMO is likely due to the duration of the printing at room temperature as well as the overexposure of pMMO to UV during curing. However, the cubic lattices retained about 85% of the enzyme based on the solid volume of the lattice (23 mm³) corresponding to the highest protein loading that was have achieved. While not wishing to be bound by any theory, it is thought that this high retention was likely due to higher cross-linking efficiency.

Since the lattice geometry did not permit precise tuning of surface area to volume ratios, due to bending of lattice struts under water surface tension, a different PμSL tool designed to generate larger parts was used to print solid and hollow PEG-pMMO cylinders with surface area to volume ratios ranging from 1.47-2.33 and diameters ranging from of 1-5 mm. The hollow tube geometry may allow more facile diffusion of reactants because both the inner and outer surfaces of the cylindrical materials would be exposed. The total print time for an array of cylinders using the large-area PμSL tool was significantly reduced to ˜1 min by eliminating z-axis resolution, and the pMMO concentration was reduced to 2.3 mg/ml to allow UV light penetration through the 1.5-3 mm depth of the resin. Remarkably, the activity of pMMO in the hydrogels increased with greater surface area to volume ratios as shown in FIG. 13 , with the highest ratio of 2.33 resulting in an average activity of 128+/−14 nmol MeOH min⁻¹mg⁻¹per cylinder, which corresponds to the highest reported physiological activity of membrane bound pMMO. The cylinders of the lowest ratio, 1.47, had an average pMMO activity of 67+/−3 nmol MeOH min⁻¹mg⁻¹. It should also be noted that the cylinders with the lowest surface area to volume ratio were only 1.5 mm in height and therefore completely submerged in the liquid phase during the activity assay, whereas all other cylinders tested were 3 mm in height and only partially submerged during the assay. Hydrogels protruding from the liquid allowed a direct interface between the gas phase and PEG-pMMO. This exposed interface likely increased the methane concentration in the PEG-pMMO material since the solubility of methane in PEG is several times higher than that in water. On average, 38% of the protein was encapsulated, although it was variable depending on the dimensions of each cylinder (27-54%). These results, combined with the results from the continuous flow reactor, indicate that an optimal pMMO material design may be hierarchical, with the smallest feature sizes at the micron scale.

Specific Methods

a. Materials

Reagents for buffers (PIPES, NaCl, and NaOH), HPLC grade methanol (≥99.9% purity), polyethylene glycol diacrylate 575 (PEGDA 575), and the cross-linking initiator, 2-hydroxy-2-methylpropiophenone (Irgacure® 1173), was purchased from Sigma-Aldrich (St. Louis, Mo.). All reagents were used as received. Methane gas (99.9% purity) was obtained from Matheson Tri-gas, Inc. (Basking Ridge, N.J.). pMMO concentrations were measured using the DC™ protein assay purchased from Bio-Rad (Hercules, Calif.). Lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) photoinitiator was synthesized following a procedure known in the art.

b. pMMO: Cell Growth and Membrane Isolation

Methylococcus capsulatus (Bath) cells were grown in 12-15 L fermentations. M. capsulatus (Bath) cells were grown in nitrate mineral salts medium (0.2% w/v KNO3, 0.1% w/v MgSO₄·7H₂O and 0.001% w/v CaCl₂·2H₂O) and 3.9 mM phosphate buffer, pH _(6.8), supplemented with 50 μM CuSO₄·5H₂O, 80 μM NaFe(III) EDTA, 1 μM Na₂MoO₄·2H₂O and trace metals solution. Cells were cultured with a 4:1 air/methane ratio at 45° C. and 300 rpm. Cells were harvested when the A₆₀₀ reached 5.0-8.0 by centrifugation at 5000×g for 10 min. Cells were then washed once with 25 mM PIPES, pH 6.8 before freezing in liquid nitrogen and storing at −80° C. Frozen cell pellets were thawed in 25 mM PIPES, pH 7.2, 250 mM NaCl buffer (herein referred to as pMMO buffer) and lysed by microfluidizer at a constant pressure of 180 psi. Cell debris was then removed by centrifugation at 20,000-24,000×g for one hr. The membrane fraction was pelleted by centrifugation at 125,000×g for one hour and washed 3 times with pMMO buffer before freezing in liquid nitrogen and storing at —80° C. Final protein concentrations were measured using the Bio-Rad DCTM assay. Typical storage concentrations ranged from 20-35 mg/ml.

c. Formation of the PEG-pMMO Hydrogels

Prior to preparation of the PEG-pMMO hydrogels, frozen as-isolated crude membranes from M. Capsulatus (Bath) (herein referred to as membrane-bound pMMO) was thawed at room temperature and used within 5 hours of thawing. Thawed membrane-bound pMMO (50-500 μg) was then mixed with PEGDA 575 in pMMO buffer at room temperature to form liquid PEG and pMMO suspensions having a final volume of 50 μl and 10-80 (v/v %) PEGDA 575. A photoinitiator (not shown in FIG. 9 ) was included in the suspension at 0.5 vol % with respect to PEGDA 575. The suspension was mixed by pipetting to homogeneity and then transferred to a 1 ml syringe with the tip removed. The syringe was then immediately placed under UV light at 365 nm, 2.5 mW/cm² intensity, for 3 min. After the UV exposure, the 50 μl PEG-pMMO hydrogel block was slowly pushed out of the syringe onto a kimwipe where it was gently blotted and then rinsed twice in pMMO buffer to remove unreacted reagents.

d. Activity Assay

All reactions were carried out in 2 ml glass reaction vials in pMMO buffer with 6 mM NADH as a reducing agent. Vials with 50-500 μg pMMO in 125 μl buffer solution were used as controls. For the immobilized enzyme samples, each 50 μl PEG-pMMO hydrogel block was placed in a vial and partially submerged in 75 μl buffer solution immediately after curing and rinsing. 1 ml of headspace gas was removed from each vial using a 2 ml gas tight glass syringe and replaced with 1 ml of methane, then the reaction vial was immediately placed in a heating block set at 45° C. and incubated for 4 min at 200 rpm. After 4 min, the samples were heat inactivated at 80° C. for 10 min. Samples were then cooled on ice for 20 min and pMMO control vials were centrifuged to remove the insoluble membrane fraction. For the cyclic activity assays using the PEG-pMMO immobilized enzyme, the reaction was stopped by opening and degassing the head space and immediately removing the solution for GC analysis. The block was then rinsed three times with 1 ml of pMMO buffer per wash and the assay was repeated. The amount of methanol generated during the reaction was measured by gas chromatography/mass spectrometry (GC/MS) analysis using an Agilent Pora-PLOT Q column and calibration curves were generated from methanol standards.

e. pMMO Flow Reactor

A simple cubic polydimethyl siloxane (PDMS) lattice with 250 micron struts and 250 micron spacing was printed using Direct Ink Write as described to provide methane permeability throughout the PEG material and to provide mechanical support. A top layer of 50 micron thick PDMS was fabricated by spin-coating Dow Corning SE-1700 PDMS diluted in toluene on a hydrophobized silicon wafer. This thin PDMS membrane prevented leakage of liquid through the membrane but provided gas permeability. Two different flow cell geometries were fabricated using polycarbonate plastic: a flow cell for a higher surface area, thin lattice (1.25 cm wide by 3 cm long) and a lower surface area, thick lattice, 1.25 by 1.25 cm. The thin lattice was 6 layers thick, and the thick lattice had 16 layers. The lattices were made hydrophilic by treating them in air plasma for 5 minutes followed by storage in deionized water. To incorporate the pMMO into the lattices, a 10 vol % concentration of PEGDA 575 was mixed with crude pMMO membrane preparations to a final concentration of 5 mg/ml pMMO. Two hundred microliters of the pMMO/PEGDA mixture were pipetted into the lattice and cured with 365 nm UV light at 2.5 mW/cm² intensity for 4 min, forming the mixed polymer (PEG/PDMS) membrane. The final concentration of pMMO in the lattices was calculated, rather than directly quantified using a protein assay, due to difficulties in quantifying the material in the lattice. The membrane was then loaded into the cell and rinsed with buffer to remove any unpolymerized material. The flow cell was placed on a hot plate calibrated with thermocouple so that the membrane would reach either 25 or 45 degrees ° C. An NADH/buffer solution (4 mg/ml NADH in PIPES pH 7.2) was prepared as the liquid phase in a 5 ml syringe, and the gas phase was prepared as 50% methane and 50% air loaded into a gas-tight 50 ml syringe. The syringes were loaded into Harvard Apparatus syringe pumps and the gas and liquid were delivered at 0.5 and 0.75 ml per hour, respectively. The gas outlet tubing was kept under 2 cm water pressure during the reaction. Fractions of liquid were collected into GC/MS autosampler vials that were kept on ice to reduce methanol evaporation and were analyzed against MeOH standards using GC/MS as described above. Methanol contamination was present in the NADH/buffer solutions, and this concentration was subtracted from the total detected in each fraction by GC/MS. No methanol contamination was found in the water used to store the PDMS. The data shown in FIG. 12B represent cumulative methanol (where the quantity of methanol produced in each fraction was added to the previous samples). Each experiment was done in triplicate; the error bars represent a standard deviation.

f 3D Printing of PEG-pMMO Hydrogels

The printing resin was prepared with 20 vol % PEGDA 575, 10 mg/ml LAP initiator, and 2.3-5 mg/ml crude pMMO in buffer. Using projection microstereolithography (POL), hydrogel blocks were printed in a cubic lattice with 100 urn open channels spaced 100 um apart and size dimensions from 1-3 mm. Solid and hollow cylinders of the same resin formulation were printed using the large area PμSL (LA PμSL) system. The cylinders had an inner diameter of 1-2.5 mm, an outer diameter of 3-5 mm, and were 1.5-3 mm high. The resin was cured with a 395 nm diode with both PμSL and LA PμSL but the intensity and exposure time varied between the systems, ranging from 11.3-20 W/cm² and 15-30 seconds per layer, respectively. Resin and printed hydrogels were stored on ice before and after the printing process. The pMMO activity assay was carried out as described above at 45° C. for 4 minutes. The methanol concentration of the activity assay and protein content of the printed hydrogels were measured as described above.

Microcapsule Embodiments

The foregoing descriptions primarily reference block polymer and/or copolymer networks as the structural arrangement most suitable for carrying out the respective chemical reactions. Those having ordinary skill in the art will appreciate that in particular applications the use of a biocatalytic microcapsule provides additional advantages and/or functionalities beyond those described above regarding polymer/copolymer networks.

However, it should be noted that the following descriptions of microcapsule embodiments may employ any of the foregoing compositions, structures, techniques, etc. described with reference to block polymer and/or copolymer networks. In some approaches for example the polymeric shell of the microcapsules may be or comprise a block polymer/copolymer network, which may comprise any combination of suitable polymers as described above with reference to block polymer/copolymer embodiments. In other approaches, the microcapsules may be embedded in a polymeric network as described hereinabove.

Utilizing biocatalytic microcapsules as described in further detail below enables production of highly efficient gas conversion systems. The presently disclosed biocatalytic microcapsules may be configured in packed stationary or moving beds, as well as fluidic beds, or embedded in meshes or adhesive compositions, allowing a broad applicability to many industrial uses requiring gas conversion (such as greenhouse gas emission reductions, food preservation, etc.).

More specifically, by encapsulating all necessary reagents/components to carry out gas conversion in a robust polymeric shell permeable to the gas species targeted for conversion, available surface area for adsorbing the target gases may be dramatically increased (e.g. on the order of approximately ten-fold relative to conventional stir tank approaches). Improvements to mass transport across the polymeric shell relative to solvation of the gases in the conventional liquid medium further improve the efficiency of gas conversion. In addition, since the biocatalyst is isolated in the interior of the polymeric shell, and includes all necessary components/reagents for renewable catalysis, the biocatalytic microcapsules of the presently disclosed inventive concepts provide improved capacity for reuse and longevity of the catalyst in its intended application.

Further still, and also an advantageous feature of using catalysts isolated in polymeric shells, the presently described inventive microcapsules may be employed in dry applications, and are therefore suitable for use as preservatives of materials sensitive to particular gases (e.g. food items are generally sensitive to ethylene and decompose more rapidly in the presence of ethylene, by converting ethylene to e.g. ethylene oxide the ethylene-driven acceleration of the decomposition process may be retarded or terminated). Accordingly, biocatalytic microcapsules as described herein allow the use of efficient gas conversion catalysts in passive environments and applications and do not require a liquid medium to facilitate mass transport of the target gases for conversion.

Further still, by selecting appropriate polymer(s) for use in the polymeric shell, preferred embodiments of biocatalytic microcapsules intended for use in passive applications such as food/medicine preservation may be non-toxic (i.e. at least to human biology). Moreover, the presently disclosed inventive microcapsules may be fabricated in a much more cost-efficient manner than traditional materials employed for such passive applications (e.g. palladium, platinum, and other precious metal catalysts, which typically are both expensive and toxic).

Accordingly, the presently disclosed inventive biocatalytic microcapsules represent a significant improvement to the function of conventional gas conversion technology, as well as an extension of gas conversion to applications and environments that are not possible using conventional techniques and technology.

These improvements are conveyed by two important, indeed critical components of the presently described inventive concepts, and include 1) a biocatalytic material, e.g. dried and reconstituted cells of suitable organisms such as methanotrophs (or components thereof such as described in greater detail below), which readily oxidize C 1-C3 gases such as methane, carbon monoxide, carbon dioxide, ethane, ethylene; propane; and/or propylene; combined with 2) the encapsulation of this material in a polymer shell which readily allows penetration of the gas phase reactants, and release of converted products. The size scale of these capsules may have a diameter in a range from approximately 10 to approximately 1000 microns, with the polymeric shell having a thickness in a range from approximately 5 microns to approximately 100 microns, in various embodiments and depending on the application to which the microcapsules are to be employed. Preferably, the capsules are spherical and the polymer shell is either hydrophobic and gas permeable, for example silicone, or amphiphilic, allowing transport of gasses and charged species, e.g. following the configuration of a block copolymer such as described hereinabove.

The biocatalytic capsules can be used in a packed bed, moving bed, or fluidized bed configuration. Alternatively, they can be immobilized or embedded in a second material, like a mesh, provided in a packet encapsulating the microcapsules but permitting gases to permeate the packet (e.g. such as packets included in certain food packaging to prevent oxidation of food products contained therein) or added to a liquid adhesive, to create a conformal coating, e.g. on a surface or as an insert in a produce box to remove ethylene. In accordance with one embodiment, the inventors have demonstrated the synthesis of stable, catalytically active microcapsules in the laboratory that oxidize propylene under ambient conditions.

Several details will now be presented regarding the structure and composition of the inventive biocatalytic microcapsules with reference to FIGS. 14A-14B, according to a preferred embodiment.

As shown in FIG. 14A, a microcapsule 1400 includes a polymeric shell 1402 defining an internal region 1404. The polymeric shell has a thickness tin a range from approximately 5 microns to approximately 100 microns, and an outer diameter d in a range from approximately 10 microns to approximately 1000 microns. As noted above the particular thickness and diameter of the microcapsule 1400 may be chosen based on the particular application to which the microcapsules will be employed.

For instance, smaller microcapsules with thinner walls may be desired in certain applications where maximum surface area and mass transport are desired, but mechanical strength is less important (e.g. in passive applications where the microcapsules are not subject to agitation or other mechanical stress, including packed, dry beds). In other embodiments, e.g. food preservation, where the microcapsules may be subject to mechanical stress associated with handling of the food items and/or packaging the thickness of the walls may be on the higher end of the spectrum noted above. Skilled artisans will appreciate, upon reading the instant descriptions, the particular thickness and/or microcapsule diameter ranges appropriate for various applications.

Moreover, the polymeric shell 1402 is preferably permeable to the target gas(es) to be catalyzed/converted by the biocatalytic components included in the microcapsule 1400. In various embodiments, polymeric shell 1402 is insoluble in aqueous solutions, and is preferably either hydrophobic and permeable to said target gases, or amphiphilic and conducts charged species and target gases across the polymeric shell 1402, which acts effectively as a selective membrane to facilitate mass transport of the target gases from the environment to the interior region 1404 of the microcapsule 1400.

Even more preferably, the polymeric shell 1402 is also permeable and/or conducts the products of catalyzing/converting the target gases. Optionally, the permeability/conductivity of the polymeric shell 1402 to the products of the catalysis/conversion reaction may be tuned by adjusting a (preferably reversible) parameter such as environmental temperature, pH, etc. to provide selective ability to release the reaction products under desired conditions.

For instance, target gases may be captured and converted at atmospheric conditions, but to avoid contaminating the environment with catalysis/conversion products the polymeric shell 1402 may be substantially (e.g. 90% or greater) impermeable to the products under atmospheric conditions. However, by elevating the temperature of the microcapsules 1400, the products may be released. Such a scheme allows isolation of the microcapsules having the catalysis/conversion products disposed therein (e.g. in a containment or collection system/facility) and selective release of such products so as to avoid simply reintroducing e.g. a reversible product which may be converted back into the undesired target gas in the operational environment.

In various approaches, the polymeric shell 1402 may include crosslinked polymers formed from one or more polymer precursors. Cros slinking and formation of the polymeric shell will be discussed in further detail below regarding FIG. 15 . In various embodiments, the polymer precursors may include any one or more of the following: polydimethylsiloxane (PDMS); polyethylene glycol (PEG); polyethylene glycol diacrylate (PEGDA); hexanediol diacrylate (HDDA); polyvinyl alcohol (PVA); poly(lactic acid) (PLA); polyimide; poly(2-methyl-2-oxazoline) (PMOXA); poly(ether ether ketone) (PEEK); cellulose acetate; polypropylene (PP); 3-[tris(trimethylsiloxy)silyl]propyl methacrylate; trimethylolpropane trimethacrylate; 2-Hydroxy-2-methylpropiophenone; and silicone acrylate.

In one preferred approach, the polymeric precursors include a mixture of the 3-[tris(trimethylsiloxy)silyl]propyl methacrylate; the trimethylolpropane trimethacrylate; and the 2-Hydroxy-2-methylpropiophenone. More preferably, the mixture includes 3-[tris(trimethylsiloxy)silyl]propyl methacrylate present in an amount from about 70 wt % to about 90 wt %; trimethylolpropane trimethacrylate present in an amount from about 10 wt % to about 30 wt %; and 2-Hydroxy-2-methylpropiophenone present in an amount from about 0.1 wt % to about 10 wt %.

In a particularly preferred embodiment, the polymer precursor mixture comprises 79.5 wt % 3-[tris(trimethylsiloxy)silyl]propyl methacrylate (preferably containing mono methyl ether hydroquinone (MEHQ) in an amount ranging from about 200-800 PPM as stabilizer, 98% pure or greater), 19.5 wt % trimethylolpropane trimethacrylate (preferably containing 250 ppm monomethyl ether hydroquinone as an inhibitor, technical grade); and 1 wt % 2-hydroxy-2-methylpropiophenone (97% pure or greater). Utilizing this mixture the inventors were able to reliably fabricate biocatalytic microcapsules 1400 having a diameter of approximately 300 microns, as shown in FIG. 14B according to one embodiment.

With continuing reference to FIG. 14A, microcapsule 1400 also includes one or more biocatalysts contained in the interior region 1404 of the polymeric shell 1402. Although the embodiment of FIG. 14A depicts a plurality of distinct types of biocatalysts, including enzymes 1408, enzyme cofactors 1410 (which may or may not be associated with enzymes); cell membrane fragments 1412, reconstituted whole cells 1414, and cytosolic components 1416, it should be understood that in various approaches microcapsules 1400 may include any one of the foregoing, or any combination thereof without departing from the scope of the presently disclosed inventive concepts. Example biocatalysts include: membrane fractions, cytosolic components, whole dried/reconstituted, and/or live cells such as soil microbes Methanococcus Capsulatus, and Ralstonia Eutropha, or the acetogenic anaerobes such as Acetobacterium woodii. Cofactors may include any combination of formate, NADH, duroquinol, etc. as would be appreciated by a person having ordinary skill in the art upon reading the present disclosures. Additionally, in some embodiments reducing agents can be supplied inside the capsule, the reducing agents being derived from proteins such as hydrogenase (which advantageously regenerate reducing equivalents with the addition of hydrogen) or methanol dehydrogenase (which regenerate reducing equivalents from the conversion of methanol product to formaldehyde) and/or formate dehydrogenase.

Preferably, the biocatalytic components included in the interior region 1404 of microcapsule 1400 include at least an entire proteome of one or more organism(s) adapted to catalyze or convert target gases for the application to which the microcapsules 1400 are to be employed, or at least those portions of a proteome of the organism that include proteins involved in the catalysis or conversion of such target gases.

The biocatalytic components 1408-1416 may be suspended in an aqueous buffer solution 1406 also disposed in the interior region of the microcapsule. Preferably, the buffer includes a reducing agent, but as noted above one advantage of using reconstituted whole cells is the lack of a need to include additional or separate reducing agent. The reconstituted whole cells provide the requisite reducing agent, if any, improving the longevity of the biocatalyst included in the microcapsule 1400. In one embodiment, the reducing agent comprises formate, and may be included in the buffer in an amount ranging from about 1 to about 100 millimolar.

Regardless of whether the buffer includes additional reducing agent, the buffer solution is preferably aqueous and has a pH in a range from approximately 4.0 to approximately 10.0.

The use of reconstituted whole cells (or, in various embodiments select subsets of cell components, such as an entire proteome of a particular organism; select enzymes and/or associated cofactors, cell membrane fragments and associated proteins; cytosolic cell components such as cytosolic proteins, organelles, etc.) encapsulated in a polymeric shell conveys several advantages in the context of the presently described inventive microcapsules.

First, such microcapsules include all necessary reagents and/or components to carry out the conversion reaction, and may not require the addition of any reducing agent to the microcapsule (typically, a reducing agent is necessary to provide renewable catalytic activity) since the reconstituted organism may advantageously include an appropriate reducing agent naturally.

Second, encapsulating such biocatalysts in a polymeric shell, e.g. as opposed to a lipid bilayer encapsulating a liposome or enzyme mixture per to conventional approaches, provides drastically improved stability of the biocatalyst and broader applicability to the biocatalyst (e.g. beyond stir-tank embodiments and including dry and/or passive applications, as well as high throughput configurations such as fluidized beds, in which lipid bilayer-based configurations would collapse).

To leverage the most efficient catalytic pathways for gas conversion, embodiments of the presently disclosed inventive concepts preferably utilize reconstituted whole cells of an organism adapted to carrying out the conversion reaction. In the context of C1-C3 gases, e.g. methane, ethane, ethylene, propylene, etc. as discussed herein, methanotrophic organisms are a generally suitable class of organism for conducting gas conversion.

The cells may be lyopholized and reconstituted in an appropriate (preferably aqueous) buffer to form a suspension of the biocatalyst(s). This suspension may be emulsified in a mixture of polymer precursors, which are in turn emulsified in an appropriate aqueous carrier fluid, and the polymer precursors may be cured to form the polymeric shell having the biocatalyst suspension disposed therein. Formation of microcapsules will be described in further detail below with reference to FIGS. 15-16 .

Turning now to FIG. 15 , a simplified top-down schematic of an exemplary apparatus 1500 for producing biocatalytic microcapsules is shown, according to one embodiment. The apparatus 1500 includes a bath 1502 filled with an aqueous carrier fluid 1504 into which droplets 1508 of a microcapsule precursor material are formed/extruded/deposited using an appropriate droplet generator 1506 of any type known in the art that would be appreciated by a skilled artisan as suitable for generating droplets 1508 having characteristics as described herein. In various embodiments, such characteristics of droplets 1508 may include, but are not limited to: emulsification structure (including but not limited to a double emulsion structure between aqueous carrier fluid 1504, polymeric precursors, and biocatalytic components in the droplets 1508, and thickness of the polymer precursor phase), composition of polymer precursors and biocatalytic components, and droplet size (e.g. diameter).

In preferred approaches, each of the droplets 1508 are deposited/extruded, etc. into the aqueous carrier fluid 1504 in the form of an emulsion, more specifically a suspension of one or more biocatalytic components (e.g. 1408-1416 as shown in FIG. 14A), the biocatalytic component suspension being emulsified in a polymer precursor layer comprising one or more polymer precursors.

As noted above, and in various embodiments, the polymer precursors may include any one or more of the following: polydimethylsiloxane (PDMS); polyethylene glycol (PEG); polyethylene glycol diacrylate (PEGDA); hexanediol diacrylate (HDDA); polyvinyl alcohol (PVA); poly(lactic acid) (PLA); polyimide; poly(2-methyl-2-oxazoline) (PXMOA); poly(ether ether ketone) (PEEK); cellulose acetate;

-   -   polypropylene (PP); 3-[tris(trimethylsiloxy)silyl]propyl         methacrylate; trimethylolpropane trimethacrylate;         2-Hydroxy-2-methylpropiophenone; and silicone acrylate.

The polymer precursors may optionally include stabilizers such as mono methyl ether hydroquinone (MEHQ), etc. to facilitate proper fluid dynamics during deposition/extrusion of droplets 1508 including but not limited to viscosity, shear, flow rate, surface tension, etc. and thus preserve the emulsion between the polymer precursor layer and the biocatalytic components.

The polymer precursors may additionally and/or alternatively comprise one or more inhibitors, e.g. photoinhibitors configured to prevent spontaneous crosslinking of polymer precursors in response to exposure to ambient light. The inhibitors are preferably selected based on the curing process and conditions to be employed to convert the polymer precursors to a polymeric shell.

In one embodiment, polymer precursor fluid includes: approximately 79.5 wt % 3-[Tris(trimethylsiloxy)silyl]propyl methacrylate (optionally but preferably including MEHQ as stabilizer, 98% purity), approximately 19.5 wt % Trimethylolpropane trimethacrylate (optionally but preferably including 250 ppm monomethyl ether hydroquinone as inhibitor, technical grade), 1 wt % 2-Hydroxy-2-methylpropiophenone (97% purity). Another option is a commercially available silicone acrylate (but used for completely different purposes), e.g. TEGO RAD 2650.

In one preferred approach, the polymeric precursors include a mixture of the 3-[tris(trimethylsiloxy)silyl]propyl methacrylate; the trimethylolpropane trimethacrylate; and the 2-Hydroxy-2-methylpropiophenone. More preferably, the mixture includes 3-[tris(trimethylsiloxy)silyl]propyl methacrylate present in an amount from about 70 wt % to about 90 wt %; trimethylolpropane trimethacrylate present in an amount from about 10 wt % to about 30 wt %; and 2-Hydroxy-2-methylpropiophenone present in an amount from about 0.1 wt % to about 10 wt %.

In a particularly preferred embodiment, the polymer precursor mixture comprises 79.5 wt % 3-[tris(trimethylsiloxy)silyl]propyl methacrylate (preferably containing mono methyl ether hydroquinone (MEHQ) in an amount ranging from about 200-800 PPM as stabilizer, 98% pure or greater), 19.5 wt % trimethylolpropane trimethacrylate (preferably containing 250 ppm monomethyl ether hydroquinone as an inhibitor, technical grade); and 1 wt % 2-hydroxy-2-methylpropiophenone (97% pure or greater).

Polymer precursors of the variety mentioned above may be obtained commercially and prepared (e.g. by solvating appropriate particles in an appropriate solvent) using techniques known in the art. The polymer precursor mixture may also be combined with the biocatalyst suspension, and an emulsion thereof generated, using techniques known in the art. This emulsion may be provided to the droplet generator 1506 for delivering droplets 1508 to the aqueous precursor fluid.

Upon delivery to the aqueous carrier fluid 1504, the droplets form a double emulsion in which the biocatalyst suspension is emulsified in the polymer precursor fluid, and the polymer precursor fluid is emulsified in the aqueous carrier fluid 1504. To facilitate forming the double emulsion, in preferred approaches the aqueous carrier fluid 1504 preferably includes water present in an amount from about 50 wt % to about 60 wt %; glycerol present in an amount from about 30 wt % to about 40 wt %; and polyvinyl alcohol present in an amount from about 1 wt % to about 5 wt %. More preferably, according to one embodiment the aqueous carrier fluid 1504 comprises approximately 58 wt % water, approximately 40 wt % glycerol, and approximately 2 wt % polyvinyl alcohol. Most preferably, monomers of the PVA are independently characterized by a molecular weight in a range from approximately 13,000-23,000 g/mol, and are approximately 87-89% hydrolyzed.

The droplets 1508 travel through the bath 1502 along a prevailing flow direction and are carried toward a curing region 1512 in which the polymer precursor layer of the droplets 1508 are cured to form a polymeric shell such as polymeric shell 1402 as shown in FIG. 14A. In preferred approaches, and in accordance with the embodiment of FIG. 15 , the curing process involves photocuring the polymer precursors, e.g. via UV crosslinking. Accordingly, apparatus 1500 may include a light source 1510 configured to emit light having predetermined characteristics (e.g. a wavelength suitable to initiate crosslinking in the polymer precursors of the droplets 1508) within the curing region 1512.

Droplets flowing through the curing region 1512 are exposed to light from the light source 1510, causing the polymer precursors to crosslink and solidify, resulting in biocatalytic microcapsules 1400. The microcapsules 1400 may continue to flow along the flow direction of the bath 1502 toward an outlet of the apparatus 1500 (not shown).

In some embodiments, microcapsules 1400 may be stored in an appropriate buffer, preferably an aqueous buffer excluding target gases for subsequent catalysis/conversion. Optionally, the microcapsules may be removed from the aqueous carrier fluid 1504 and washed (e.g. in water) and/or dried for subsequent use in an embedded matrix, mesh, adhesive, etc. for dry applications such as food preservation.

Accordingly, and with reference to FIG. 16 , a preferred embodiment of a method 1600 of forming microcapsules for selective catalysis of gases is shown. The method 1600 may be performed in any suitable environment, including but not limited to the apparatus as shown in FIG. 15 , without departing from the scope of the present disclosures. Moreover, the method 1600 may include any number of additional or alternative operations than shown in FIG. 16 , in alternate embodiments, so long as the result is formation of biocatalytic microcapsules as described herein.

The present method 1600 may be implemented in conjunction with features from any other embodiment listed herein, such as those described with reference to the other FIGS. Of course, the method 1600 and others presented herein may be used in various applications and/or in permutations, which may or may not be specifically described in the illustrative embodiments listed herein.

Returning now to FIG. 16 , in accordance therewith method 1600 includes operation 1602, in which at least one biocatalyst is emulsified in a polymer precursor mixture. The emulsion may be prepared using any suitable technique that would be appreciated as suitable by a skilled artisan after reading the instant descriptions. Preferably, the biocatalyst(s) is/are suspended in an appropriate buffer fluid, which may or may not include reducing agent, and is preferably an aqueous buffer. The biocatalyst suspension may include any number or type of biocatalytic components as described herein, e.g. with reference to FIG. 14A above. The polymer precursor mixture may have any composition as described hereinabove, or equivalents thereof that would be appreciated by skilled artisans upon reading the present disclosure.

In operation 1604, method 1600 involves emulsifying the polymer precursor mixture in an aqueous carrier solution. The process of emulsifying the polymer precursor mixture, which is preferably present in the form of a droplet such as droplets 1508 as shown in FIG. 15 , involves controlling fluid dynamics within a suitable bath containing the aqueous carrier solution into which the droplets (having the polymer precursor mixture disposed in an outer layer thereof) are formed/deposited. In addition, the composition of the aqueous carrier solution and the polymer precursor mixture should be formulated so that an emulsion therebetween may be formed. Accordingly, the polymer precursor mixture and aqueous carrier solution may have a composition as described elsewhere herein, according to various embodiments.

With continuing reference to FIG. 16 , in operation 1606 of method 1600 polymer precursor(s) of the polymer precursor mixture are cured, e.g. via crosslinking, to form a polymeric shell therefrom. This results in a plurality of microcapsules each independently comprising: a polymeric shell permeable to one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell, e.g. in accordance with microcapsules 1400 as shown in FIGS. 14A-14B. Preferably, the curing involves projection micro stereolithography.

Applications/Uses

Embodiments of the present invention may be used in a wide variety of applications, and potentially any industrial application requiring more efficient and higher-throughput use of enzymes to catalyze chemical reactions. Illustrative applications in which embodiments of the present invention may be used include, but are not limited to, fuel conversion (e.g., natural gas to liquid fuel), chemical production, pharmaceutical production, and other processes where a chemical conversion is catalyzed by enzymes, especially at phase boundaries (e.g., reaction involving a gas and a liquid, polar and non-polar species, aqueous and non-aqueous species, etc.).

In accordance with several embodiments, and with reference to method 1700 as shown in FIG. 17 , the presently disclosed inventive biocatalytic microcapsules may be utilized in techniques for catalyzing one or more target gases. The method 1700 may be performed in any suitable environment, including but not limited to stir tanks, fluidized beds, packed beds, moving beds, embedded meshes and/or adhesives, etc., without departing from the scope of the present disclosures. Moreover, the method 1700 may include any number of additional or alternative operations than shown in FIG. 17 , in alternate embodiments, so long as the result is catalysis/conversion of target gases (e.g. C1-C3 gases) using biocatalytic microcapsules as described herein.

In one embodiment, method 1700 includes exposing a plurality of the biocatalytic microcapsules to the one or more target gases in operation 702. Importantly, the biocatalytic microcapsules each independently comprise: a polymeric shell permeable to the one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell.

The exposure may be performed in any suitable manner, such as passively allowing target gases to flow over or through the microcapsules (which may optionally be arranged in a bed, mesh, adhesive, etc.) or actively passing target gases through the microcapsules (e.g. driven by pressure, via bubbling gases through a fluidized bed, agitation of the microcapsules, movement of microcapsules within a bed or use of a moving bed, etc.) depending on the application in question.

For example, for a carbon dioxide capture application packed beds and fluidized beds are most viable. For smaller scale industrial applications such as ethylene or methane conversion fixed beds, or packed beds are preferred. Generally, avoiding moving parts for smaller applications is desirable to minimize maintenance and energy consumption, while for larger scale applications a fluidized bed is a preferred, energy efficient configuration. For applications such as food and medicine preservation, involving e.g. capture of ethylene, dry, small scale configurations such as microcapsules embedded in meshes or adhesives are desired to facilitate use of the microcapsules in the packaging containing the food or medicine to be preserved. Of course, in various applications different configurations of the microcapsules may be employed without departing from the scope of the present disclosures.

The target gases are preferably C1-C3 compounds, e.g. methane, carbon monoxide, carbon dioxide, ethane, ethylene; propane; and/or propylene, in various embodiments, and are preferably catalyzed by one or more biocatalytic components selected from: one or more enzymes configured to catalyze the one or more target gases; one or more enzyme cofactors; one or more cell membrane fragments; one or more cytosolic cell components; and reconstituted whole cells.

Various embodiments herein have been described with reference to catalysis or conversion of target gases using biocatalysts. It should be understood that catalysis, conversion, etc. of target gases may include any suitable chemical modification such as reduction or oxidation of the target gases into another species (gas or liquid, preferably). In more embodiments, catalysis/conversion may include capturing such target gases, which may be reversibly released into a collection environment, e.g. in the case of carbon dioxide. Skilled artisans reading the present disclosures will appreciate other equivalent suitable forms of catalysis and/or conversion of target gases to which the presently described inventive microcapsules may be applied.

It should be noted that methodology presented herein for at least some of the various embodiments may be implemented, in whole or in part, in computer hardware, software, by hand, using specialty equipment, etc. and combinations thereof.

Moreover, any of the structures and/or steps may be implemented using known materials and/or techniques, as would become apparent to one skilled in the art upon reading the present specification.

The inventive concepts disclosed herein have been presented by way of example to illustrate the myriad features thereof in a plurality of illustrative scenarios, embodiments, and/or implementations. It should be appreciated that the concepts generally disclosed are to be considered as modular, and may be implemented in any combination, permutation, or synthesis thereof. In addition, any modification, alteration, or equivalent of the presently disclosed features, functions, and concepts that would be appreciated by a person having ordinary skill in the art upon reading the instant descriptions should also be considered within the scope of this disclosure.

While various embodiments have been described above, it should be understood that they have been presented by way of example only, and not limitation. Thus, the breadth and scope of an embodiment of the present invention should not be limited by any of the above-described exemplary embodiments, but should be defined only in accordance with the following claims and their equivalents. 

What is claimed is:
 1. A method of forming microcapsules for selective catalysis of gases, the method comprising: emulsifying at least one biocatalyst in a polymer precursor mixture; emulsifying the polymer precursor mixture in an aqueous carrier solution; crosslinking one or more polymer precursors of the polymer precursor mixture to form a plurality of microcapsules each independently comprising: a polymeric shell permeable to one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell.
 2. The method as recited in claim 1, wherein emulsifying at least one biocatalyst in a polymer precursor mixture and emulsifying the polymer precursor mixture in an aqueous carrier solution forms a double emulsion comprising the biocatalyst, the polymer precursor mixture, and the aqueous carrier solution.
 3. The method as recited in claim 1, wherein the biocatalyst is suspended in a buffer.
 4. The method as recited in claim 3, wherein the buffer excludes reducing agents.
 5. The method as recited in claim 1, the at least one biocatalyst comprising one or more biocatalytic components selected from: one or more enzymes configured to catalyze the one or more target gases; one or more enzyme cofactors; one or more cell membrane fragments; one or more cytosolic cell components; and reconstituted whole cells.
 6. The method as recited in claim 1, wherein the aqueous carrier solution comprises: water present in an amount from about 50 wt % to about 60 wt %; glycerol present in an amount from about 30 wt % to about 40 wt %; and polyvinyl alcohol present in an amount from about 1 wt % to about 5 wt %.
 7. The method as recited in claim 1, wherein the polymer precursor mixture comprises one or more polymer precursors selected from a group consisting of: polydimethylsiloxane (PDMS); polyethylene glycol (PEG); polyethylene glycol diacrylate (PEGDA); hexanediol diacrylate (HDDA); polyvinyl alcohol (PVA); poly(lactic acid) (PLA); polyimide; poly(2-methyl-2-oxazoline) (PXMOA); poly(ether ether ketone) (PEEK); cellulose acetate; polypropylene (PP); and silicone acrylate.
 8. The method as recited in claim 1, comprising: isolating the plurality of microcapsules from the aqueous carrier solution; and drying the plurality of microcapsules.
 9. A method for catalyzing one or more target gases using biocatalytic microcapsules, the method comprising: exposing a plurality of the biocatalytic microcapsules to the one or more target gases, wherein the biocatalytic microcapsules each independently comprise: a polymeric shell permeable to the one or more target gases; and at least one biocatalyst disposed in an interior of the polymeric shell.
 10. The method as recited in claim 9, wherein the biocatalytic microcapsules are arranged in a configuration selected from: a packed bed; a moving bed; a fluidized bed; immobilized in or on a substrate; a microcapsule-embedded mesh; a packet containing the biocatalyic microcapsules; and a microcapsule-embedded adhesive.
 11. The method as recited in claim 9, the one or more target gases each independently being a C1-C3 compound.
 12. The method as recited in claim 11, the one or more target gases being selected from: methane, carbon monoxide, carbon dioxide, ethane, ethylene; propane; and propylene, hydrogen and oxygen.
 13. The method as recited in claim 9, the at least one biocatalyst comprising one or more biocatalytic components selected from: one or more enzymes configured to catalyze the one or more target gases; one or more enzyme cofactors; one or more cell membrane fragments; one or more cytosolic cell components; and reconstituted whole cells. 